This research was supported by National Institutes of Health grant (R01-DE025985) and by an American Cancer Society Research Scholar Award (RSG-13-383-MPC). This work was also supported by grants from the U.S. Department of Agriculture (2015-67015-23268 and 2014-67015-21787) and New Research Initiative Grant from University of California, Davis.

CAUTION: Cell lines used in this procedure contain infectious viruses, be careful and only proceed in level 2 BSL facility or higher.
1. Cell Preparation and Maintenance
<ol>
	<li>Culture TREx-K-RTA BCBL-1 cells (or another KSHV infected PEL cell lines) in RPMI1640 medium supplemented with 15% fetal bovine serum (FBS) and 1% penicillin streptomycin glutamine solution.</li>
	<li>Grow cells at 37 &#176;C with 5% carbon dioxide (CO2), making sure to continuously grow and to split cells by a ratio of 1:4 every 2&#8211;4 days.</li>
	<li>Monitor cell cultures with a light microscope every day to confirm good cell growth and morphology, otherwise it is recommended to restart the cell culture or to adjust the cell culture conditions accordingly.</li>
</ol>
2. Thymidine Synchronization and Lytic Induction
<ol>
	<li>In a 10 cm Petri dish, plate TREx-K-Rta BCBL-1 cells in freshly prepared media (1 x 106 cells/2 mL of media)</li>
	<li>Add 200 mM thymidine (final concentration: 2 mM) to the dish, mix and incubate for 18 h.</li>
	<li>Centrifuge the cells for 5 min at 500 x g. Wash the cells with sterile 1x phosphate-buffered saline (PBS), centrifuge again, and resuspend cells in freshly prepared media. Repeat for a total of 3 washes. Resuspend in freshly prepared media.</li>
	<li>Allow the cells to exit S-phase for 8 h.</li>
	<li>Add thymidine (final concentration: 2 mM), mix and incubate for 16 h.</li>
	<li>Centrifuge the cells for 5 min at 500 x g. Wash the cells with sterile 1x phosphate-buffered saline (PBS), centrifuge again, and resuspend cells in freshly prepared media. Repeat for a total of 3 washes. Resuspend in freshly prepared media.</li>
	<li>To induce viral reactivation, add 12-O-tetradecanoyl-phrobol-13-acetate (TPA, final concentration 20 ng/mL) and doxycycline (DOX, final concentration 100 ng/mL) to the cell culture, mix and incubate for 4 h. For cell lines without a K-Rta inducible cassette, stimulate reactivation using TPA (final concentration 20 ng/mL and sodium butyrate (1 mM)).</li>
	<li>Centrifuge the cells for 5 min at 500 x g. Wash the cells with sterile 1x phosphate-buffered saline (PBS), centrifuge again, and resuspend cells in complete culture media. Repeat for a total of 3 washes. Resuspend in complete culture media.</li>
	<li>Allow the cells to grow for 24 h.</li>
</ol>
3. Fixation and Permeabilization
NOTE: Before proceeding, ensure fixing solution (3.7% formaldehyde in DEPC-treated PBS), diethyl pyrocarbonate-treated phosphate-buffered saline (DEPC-PBS), glycine DEPC PBS (final concentration of glycine: 100 mM) and permeabilizating solution (50% acetone, 50% methanol) are prepared. Typically, each slide will require at least 0.5 million cells, multiply if necessary and include extra cells for back up especially if cells are not healthy.
<ol>
	<li>Collect cells into a 1.5 mL microcentrifuge tube.</li>
	<li>Centrifuge the cells for 2 min at 200 x g. Wash the cells 3 times with 1 mL of sterile DEPC PBS. Resuspend in 1.2 mL of DEPC PBS.</li>
	<li>Place the appropriate number of coverslips (determined by the experimental setup) into the bottoms of a 6-well plate.</li>
	<li>Pipette 200 &#181;L of the cell DEPC PBS mixture onto each coverslip, ensure that at least 0.5 million cells are on each slide. Allow the cells to settle for 2 min, and aspirate excess DEPC PBS leaving only a thin layer of cells. It is normal for cells to be removed in this process. 
	Caution: PFA is toxic, wear appropriate protection.</li>
	<li>Carefully add 1 mL of fixing solution (3.7% formaldehyde in DEPC PBS) to each cover slip. Allow the cells to fix for 10 min.</li>
	<li>Wash coverslips 3 times with DEPC PBS.</li>
	<li>Carefully add 1 mL of glycine DEPC PBS (final concentration of glycine: 100 mM) to each cover slip. Allow the cells to quench for 5 min.</li>
	<li>Add 1.5 mL of DEPC PBS and either place on a shaker for 5 min or shake firmly but gently in hand for 1 min. Aspirate the excess DEPC PBS. Be careful not to disrupt the cells. Repeat this step for a total of 3 washes.</li>
	<li>Carefully add 1 mL of permeabilizing solution (50% acetone, 50% methanol) to each cover slip. Allow the cells to permeabilize for 15 min. 
	NOTE: At this point, the coverslips can be stored in a -20 &#176;C freezer for no longer than a week. Avoid freezing or minimize the time spent in the freezer as image quality can degrade quickly. Make sure that when in the freezer that the cells remain wet and covered in methanol/acetone, and to ensure that they properly rehydrate with DEPC PBS after removal from the freezer.</li>
	<li>Add 1.5 mL of DEPC PBS and either place on a shaker for 5 min or shake firmly but gently in hand for 1 min. Aspirate the excess DEPC PBS, be careful not to disrupt the cells. Repeat this step for a total of 3 washes.</li>
</ol>
4. IFA and RNA-FISH
NOTE: Regarding RNA FISH Probe labeling and preparation: For the K-Rta intron which is 959 base pairs, 31 different probes were generated that were 20 base pairs each, containing a GC content of about 50%. Working and stock solutions of the probes were aliquoted into 100 &#181;M and 2.5 &#181;M concentrations, respectively.
<ol>
	<li>Label microscope slides with necessary experimental information or some sort of distinguishable label.</li>
	<li>Line the bottom of a sealable plastic container with a moist paper towel.</li>
	<li>Prepare the primary antibody solution (1.5 &#181;L of LANA antibody, 0.75 &#181;L of RNA Pol II antibody, 3 &#181;L of baker&#39;s yeast tRNA, DEPC PBS up to 30 &#181;L), and multiply recipe for each coverslip present. Place 30 &#181;L of primary antibody solution onto each labeled microscope slide.
	<ol>
		<li>Titrate the primary antibody of interest first and use the optimal amount. Be sure to use purified IgG, as serum (ascites fluid) may contain large amount of RNase.</li>
	</ol>
	</li>
	<li>Place the coverslip (cells facing the solution) onto the labeled microscope slides and move the slides into the container with the moist paper towel. Be careful not to introduce bubbles. Cover the exterior of the container with plastic wrap. Move the container into an incubator set to 37 &#176;C for 1 h.</li>
	<li>Prepare a FISH wash buffer (2x saline sodium citrate buffer (SSC) (final), 10% formamide, 90% DEPC dH2O).</li>
	<li>Prepare 2x Hybridization Buffer (4x SSC, 20% dextran sulfate).</li>
	<li>Prepare the secondary antibody and FISH intron probe buffer (20 &#181;L of 2x hybridization buffer (final 1x Hyb Buffer), 4 &#181;L of baker&#39;s yeast tRNA (final 10%), 4 &#181;L of formamide (final 10%), intron K-Rta probe (final 125 nM), 0.8 &#181;g of secondary antibody, DEPC dH2O up to 40 &#181;L). Multiply recipe for every slide needed.</li>
	<li>After incubation, wash cells 3 times with DEPC PBS in a 6 well plate for 5 min each.</li>
	<li>Wash with FISH wash buffer 3 times for 5 min each. Keep coverslip in FISH-wash buffer until finish preparing mixture of secondly antibody and intron probe (step 4.7).</li>
	<li>Clean glass slides used for primary antibody incubation for hybridization.</li>
	<li>Place 40 &#181;L of the secondary antibody and FISH intron probe containing solution onto each glass slide.</li>
	<li>Place the coverslips with the cells onto the slides with the cell side facing down on top of the buffer, be careful not to introduce bubbles.</li>
	<li>Place the slides into a plastic container with a moist paper towel on the bottom. Wrap the plastic container in first plastic wrap and then aluminum foil.</li>
	<li>Incubate the container with the slides at 37 &#176;C for 16-24 h.</li>
	<li>After incubation, carefully slide the coverslip from the edge of the glass slide, and put back into 6 well plates facing up, and wash cells with FISH-wash buffer 3 times in a 6 well plate for 5 min each.</li>
	<li>Wash the cells 2 times with 2x SSC.</li>
	<li>Add DAPI (1:1,000) into the 2x SSC and allow to sit for 5 min at room temperature.</li>
	<li>Wash the cells 2 times with 2x SSC.</li>
	<li>Clean the glass slides used for hybridization, and add 10 &#181;L of mounting solution onto the glass slides.</li>
	<li>Place the coverslips face down onto the mounting solution. Be careful not to introduce bubbles.</li>
	<li>With a laboratory tissue, gently remove the excess mounting solution, being careful not to squash the cells.</li>
	<li>Using nail polish, apply an ample layer around the edge of the coverslips and allow to dry.</li>
	<li>Proceed to perform fluorescence microscopy.</li>
</ol>
5. 3D Fluorescence Microscopy
NOTE: While protocols will vary with the type of fluorescence microscope system used, the following steps will help to ensure the acquisition of optimal image data for quantitative analysis.
<ol>
	<li>Pretreat fixed samples and mount in anti-fade media to delay fluorescence photobleaching during imaging.</li>
	<li>Use a high quality objective (such as a 60X 1.42 N.A oil-immersion lens), which can capture a whole cell (or cell nucleus) in the field-of-view.</li>
	<li>Set exposure parameters (e.g., excitation power, exposure time) for each fluorescence color channel, using positive- and negative-control samples. Acquire fluorescence images. 
	NOTE: Positive control samples that contain only one fluorescent label should also be used to determine the level of &#34;crosstalk&#34; between color channels. For imaging fixed cell specimens, fluorescence excitation power can be reduced (with slightly longer exposure times) to minimize photobleaching.</li>
	<li>Once the imaging protocol has been established, maintain the same exposure parameters consistent for all samples in a study &#8212; so that image data can be accurately analyzed and compared.</li>
	<li>Perform post-acquisition image processing and 3D reconstruction. Identify the K-Rta transcripts by RNA-FISH signal (red), LANA molecules by immunofluorescence (green), and nuclear chromatin (where applicable) stained with DAPI (blue). 
	NOTE: Regions of the cell nucleus, in which K-Rta and LANA are co-clustered, are thus believed to be associated with viral transcription factories and replication complex.</li>
</ol>

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S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myc+dynamically+and+preferentially+relocates+to+a+transcription+factory+occupied+by+Igh.">Myc dynamically and preferentially relocates to a transcription factory occupied by Igh.</a> PLoS Biol. 5 (8), 192 (2007).</li><li>Fanucchi, S., Shibayama, Y., Burd, S., Weinberg, M. S., Mhlanga, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromosomal+contact+permits+transcription+between+coregulated+genes.">Chromosomal contact permits transcription between coregulated genes.</a> Cell. 155 (3), 606-620 (2013).</li><li>Borah, S., Darricarrere, N., Darnell, A., Myoung, J., Steitz, J. 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The authors of this paper have no financial ties or competing conflicts of interests.

There are some aspects of the protocol that can be altered to accommodate unusual circumstances. The choice of fixation and permeabilization buffers can also be altered. For fixation buffers, paraformaldehyde is also effective and ethanol can be used for permeabilization. 
Quenching the formaldehyde with glycine PBS is recommended as it prevents formaldehyde from disabling the antibodies in absence of bovine serum albumin (BSA), but if the coverslips are washed thoroughly enough, the glycine P...

It has become increasingly clear that the spatiotemporal organization of the nucleus plays an important role in modulating finely-tuned gene expression in eukaryotes. Most tissue specific genes are distributed over many chromosomes and need to be regulated synchronously in order to respond to specific stimuli in a coordinated manner1. Cells construct active chromatin hubs (ACH) as a way of bringing together genes and their cis-regulatory components to a specific nuclear space1.
It has also been demonstrated by various studies that although the nucleus seems very dense and viscous, biologically active molecules can traverse the nucleus rather quickly via diffusion2. As a consequence of this ephemeral property, most DNA binding proteins &#39;jump&#39; from binding site to binding site, feeling their way around the nuclear space, which allows for a highly adaptive and versatile nucleus2.
Despite this dynamic behavior of biomolecules within the nucleus, nuclear bodies without membranes such as (but not limited to) the nucleolus, Cajal bodies, and promyelocytic leukemia nuclear bodies (PML-NB) still exist. It is through a variety of mechanisms such as tandem DNA repeats (nucleolus), rRNA (nucleolus) and structural proteins such as coilin (Cajal bodies) or PML proteins (PML-NB) that hold these constructs together3,4,5. These structures and other congregations such as transcription factories serve as scaffolds that not only increase the local concentration of required components, but also regulate the composition of proteins and nucleic acids within them to ultimately create a central site for efficient cellular functions6.
Visualizing when and where nuclear structures form provides a wealth of information to researchers studying epigenetics. From a virology perspective, the reactivation of latently infected viruses, such as KSHV, significantly alters the landscape of the nucleus and distribution of nuclear enzymes to shift transcription primarily from cellular genes to viral genes, ultimately to produce fully functional viral progeny7,8. How does KSHV manipulate the cellular gene expression machinery to facilitate viral gene expression? Such information could also shed light on temporal cellular gene regulatory mechanisms.
Like all other herpesviruses, KSHV has two life cycles called lytic replication and latency. KSHV primarily resides in the latency stage, in which most of its viral gene expression is silenced, except latency associated genes9,10. During latency KSHV produces latency associated nuclear antigen (LANA), which constitutively binds the viral genomes and tethers viral chromatin to the human chromosome11. Because of LANA&#39;s intimate relationship with the viral genome, it is possible to use IFA and DAPI to stain and locate where the viral episomes were in relation to the host chromatin.
To study KSHV reactivation at the single episome level and the association with other viral episomes in an infected cell, a strategy to locate actively transcribing viral chromatin in situ has been established. Accordingly, LANA and RNAPII IFA with intron RNA-FISH were combined, by generating RNA-FISH probes that bind to the intron region (exact probe sequences can be found in the Izumiya lab&#39;s most recent publication8) of KSHV K-Rta-the key viral protein that is essential and sufficient for KSHV reactivation-it was possible to identify where transcription was actually taking place8,11,12,13,14,15. This intron RNA-FISH technique enables researchers to visualize where mRNA is being transcribed immediately before it is spliced and exported to cytoplasm16,17.
KSHV can be reactivated by various chemical stimuli including phorbol esters such as 12-O-Tetradeconoyl-phorbol-13acetate (TPA) and histone deacetylase inhibitors such as sodium butyrate, additionally KSHV can be induced to reactivate by overexpression of the viral transcription factor, K-Rta19. Researchers have successfully increased the efficiency of KSHV reactivation by synchronizing the cell&#39;s cycles prior to inducing reactivation18. Thus, for these particular studies, cells were synchronized using a double thymidine block (protocol described below) and incubated with TPA and doxycycline (Dox) for a short time. Doxycyline was used because the cell line utilized in these experiments has a doxycycline-inducible K-Rta cassette, which was cloned from cDNA and does not include the intron region of K-Rta. Although it is possible to reactivate KSHV using only induced K-Rta expression, it has been proven by other researchers that due to a variety of different biochemical factors K-Rta expression alone proves to be a feeble reactivation stimuli20. By combining all of these, and by limiting the drug incubations to a short period of time, a robust but not overly artificial KSHV reactivation was achieved for imaging.
After labeling LANA, RNAPII, K-Rta introns, and DNA as described in this paper, 3D fluorescence imaging was performed using a widefield deconvolution microscope. After processing with imaging software, the spatial distribution of active viral episomes can be properly evaluated. Using this technique, the central questions regarding the fundamental nature of the formation of active chromatin hubs and other nuclear structures can be studied. Having identical viral episomes in a single cell that processes the same regulatory elements may represent a unique research tool to deepen the understanding of spatiotemporal gene regulatory mechanisms.
A limitation of imaging multiple cell specimens fixed at different time points to characterize an inherently dynamic molecular process is that subtle or small-scale changes in fluorescence distribution are undetected or deemed insignificant. This is true unless in the rare instance, every cell observed displays the same subtle change. Thus, the full spatiotemporal relationship of active viral transcription and other nuclear structures cannot be critically evaluated using fixed imaging. To address these technical challenges, the best approach is to image live cells that have marked viral episomes and to follow the location of key cellular enzymes over time.

<table><tbody><tr><td>RPMI medium 1650 GlutaMAX</td><td>Gibco by Life Technologies</td><td>61870-036</td><td>Needs to be warmed to 37 &amp;deg;C</td></tr><tr><td>Fetal Bovine Serum</td><td>Omega Scientific Inc.</td><td>FB-11</td><td></td></tr><tr><td>Pen Strep Glutamine (100x)</td><td>Gibco by Life Technologies</td><td>10378-016</td><td></td></tr><tr><td>Thymidine&amp;nbsp;</td><td>Sigma Aldrich</td><td>T9250</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered Solution</td><td>Gibco by Life Technologies</td><td>14190-144</td><td></td></tr><tr><td>TPA</td><td>Sigma Aldrich</td><td>P1585</td><td></td></tr><tr><td>Sodium Butyrate&amp;nbsp;</td><td>Sigma Aldrich</td><td>B5887</td><td></td></tr><tr><td>Formaldehyde Solution&amp;nbsp;</td><td>Sigma Aldrich</td><td>252549</td><td>Corrosive, toxic, health hazard</td></tr><tr><td>Diethyl Pyrocarbonate</td><td>Sigma Aldrich</td><td>D-5758</td><td>Acute toxicity&amp;nbsp;</td></tr><tr><td>Glycine</td><td>Sigma Aldrich</td><td>410225</td><td></td></tr><tr><td>Acetone</td><td>Sigma Aldrich</td><td>179124</td><td>Flammable, toxic</td></tr><tr><td>Methanol</td><td>Fisher Chemical&amp;nbsp;</td><td>67-56-1</td><td>Flammable, toxic, health hazard</td></tr><tr><td>Rat monoclonal antibody to KSHV/HHV-8 ORF73</td><td>Advanced Biotechnologies</td><td>13-210-100</td><td></td></tr><tr><td>Anti-RNA Polymerase II Antibody clone CTD4H8</td><td>EMD Milipore</td><td>05-623</td><td></td></tr><tr><td>Ribonucleic acid, transfer from baker&amp;rsquo;s yeast (S. cerevisiae)&amp;nbsp;</td><td>Sigma Aldrich</td><td>9014-25-9</td><td></td></tr><tr><td>Saline sodium citrate buffer 20x (SSC)</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>15557044</td><td></td></tr><tr><td>Formamide&amp;nbsp;</td><td>Sigma Aldrich</td><td>295876</td><td></td></tr><tr><td>Omnipur Dextran Sulfate, 50% solution</td><td>EMD Milipore</td><td>1916C093</td><td></td></tr><tr><td>DAPI (4&amp;rsquo;,6-diamindino-2-phenylindole, Dihydrochloride)</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>D1306</td><td></td></tr><tr><td>slowFade Gold antifade reagent</td><td>Life Technologies</td><td>S36936</td><td></td></tr><tr><td>Micro cover glass 22x22</td><td>Matsunami</td><td>C022221</td><td></td></tr><tr><td>VoloCITY</td><td></td><td>3D imaging software</td><td></td></tr><tr><td>DeltaVision microscope</td><td></td><td></td><td></td></tr><tr><td>Superfrost/Plus microscope slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr></tbody></table>

functional imaging of viral transcription factories using 3d fluorescence microscopy

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to understand where and when transcription is taking place within nuclear space and to visualize the relationship between episomes infected within the same cell's nucleus. Here, both immunofluorescence (IFA) and RNA-FISH have been combinedto identify actively transcribing Kaposi's sarcoma-associated herpesvirus (KSHV) episomes. By staining KSHV latency-associated nuclear antigen (LANA), it is possible to locate where viral episomes exist within the nucleus. In addition, by designing RNA-FISH probes to target the intron region of a viral gene, which is expressed only during productive infection, nascent RNA transcripts can be located. Using this combination of molecular probes, it is possible to visualize the assembly of large viral transcription factories and analyze the spatial regulation of viral gene expression during KSHV reactivation. By including anti-RNA polymerase II antibody staining, one can also visualize the association between RNA polymerase II (RNAPII) aggregation and KSHV transcription during reactivation.

A slightly abridged protocol was performed where BCBL-1 cells were used and only LANA and K-Rta RNA were stained (Figure 1). This experiment allows us to examine where actively transcribing viral episomes are and the heterogeneity of response to KSHV reactivation stimuli in a population of cells. In the cell indicated by the arrow in Figure 1, the distinct K-Rta fluorescence in regions that closely match the distribution of viral...

Watch this Scientific Journal Video about Functional Imaging of Viral Transcription Factories Using 3D Fluorescence Microscopy at JoVE.com

Functional Imaging of Viral Transcription Factories Using 3D Fluorescence Microscopy

Viral transcriptional factories are discrete structures that are enriched with cellular RNA polymerase II to increase viral gene transcription during reactivation. Here, a method to locate sites of actively transcribing viral chromatin in 3D nuclear space by a combination of immunofluorescence staining and in situ RNA hybridization is described.

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to ...

functional-imaging-viral-transcription-factories-using-3d

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

7.0K Views.  University of California, Davis. Viral transcriptional factories are discrete structures that are enriched with cellular RNA polymerase II to increase viral gene transcription during reactivation. Here, a method to locate sites of actively transcribing viral chromatin in 3D nuclear space by a combination of immunofluorescence staining and in situ RNA hybridization is described.

Video: Functional Imaging of Viral Transcription Factories Using 3D Fluorescence Microscopy

Visualizing Cell-to-cell Transfer of HIV using Fluorescent Clones of HIV and Live Confocal Microscopy

By fusing the green fluorescent protein to their favorite proteins, biologists now have the ability to study living complex cellular processes using fluorescence video microscopy. To track the movements of the human immunodeficiency virus core protein during cell-to-cell transmission of human immunodeficiency virus, we have GFP-tagged the Gag protein in the context of an infectious molecular clone of HIV, called HIV Gag-iGFP. We study this viral clone using video confocal microscopy. In the following visualized experiment, we transfect a human T cell line with HIV Gag-iGFP, and we use fluorescently labeled uninfected CD4+ T cells to serve as target cells for the virus. Using the different fluorescent labels we can readily follow viral production and transport across intercellular structures called virological synapses. Simple gas permeable imaging chambers allow us to observe synapses with live confocal microscopy from minutes to days. These approaches can be used to track viral proteins as they move in from one cell to the next.

This visualized experiment is a guide for utilizing a fluorescent molecular clone of HIV for live confocal imaging experiments.

By fusing the green fluorescent protein to their favorite proteins, biologists now have the ability to study living complex cellular processes using ...

Immunology and Infection

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

Bioengineering

Live Cell Imaging of Alphaherpes Virus Anterograde Transport and Spread

Advances in live cell fluorescence microscopy techniques, as well as the construction of recombinant viral strains that express fluorescent fusion proteins have enabled real-time visualization of transport and spread of alphaherpes virus infection of neurons. The utility of novel fluorescent fusion proteins to viral membrane, tegument, and capsids, in conjunction with live cell imaging, identified viral particle assemblies undergoing transport within axons. Similar tools have been successfully employed for analyses of cell-cell spread of viral particles to quantify the number and diversity of virions transmitted between cells. Importantly, the techniques of live cell imaging of anterograde transport and spread produce a wealth of information including particle transport velocities, distributions of particles, and temporal analyses of protein localization. Alongside classical viral genetic techniques, these methodologies have provided critical insights into important mechanistic questions. In this article we describe in detail the imaging methods that were developed to answer basic questions of alphaherpes virus transport and spread.

Live cell imaging of alphaherpes virus infections enables analysis of the dynamic events of directed transport and intercellular spread. Here, we present methodologies that utilize recombinant viral strains expressing fluorescent fusion proteins to facilitate visualization of viral assemblies during infection of primary neurons.

Advances in live cell fluorescence microscopy  techniques, as well as the construction of recombinant viral strains that  express fluorescent fusion ...

Methodology for the Efficient Generation of Fluorescently Tagged Vaccinia Virus Proteins

Tagging of viral proteins with fluorescent proteins has proven an indispensable approach to furthering our understanding of virus-host interactions. Vaccinia virus (VACV), the live vaccine used in the eradication of smallpox, is particularly amenable to fluorescent live-cell microscopy owing to its large virion size and the ease with which it can be engineered at the genome level. We report here an optimized protocol for generating recombinant viruses. The minimal requirements for targeted homologous recombination during vaccinia replication were determined, which allows the simplification of construct generation. This enabled the alliance of transient dominant selection (TDS) with a fluorescent reporter and metabolic selection to provide a rapid and modular approach to fluorescently label viral proteins. By streamlining the generation of fluorescent recombinant viruses, we are able to facilitate downstream applications such as advanced imaging analysis of many aspects of the virus-host interplay that occurs during virus replication.

A rapid and modular protocol for the generation of recombinant vaccinia viruses expressing fluorescently&#160;tagged proteins simultaneously using the method of transient dominant selection is described here.

Tagging of viral proteins with fluorescent  proteins has proven an indispensable approach to furthering our understanding  of virus-host interactions. ...

Vaccinia Reporter Viruses for Quantifying Viral Function at All Stages of Gene Expression

Poxviruses are a family of double stranded DNA viruses that include active human pathogens such as monkeypox, molluscum contagiousum, and Contagalo virus. The family also includes the smallpox virus, Variola. Due to the complexity of poxvirus replication, many questions still remain regarding their gene expression strategy. In this article we describe the conceptualization and usage of recombinant vaccinia viruses that enable real-time measurement of single and multiple stages of viral gene expression in a high-throughput format. This is enabled through the use of spectrally distinct fluorescent proteins as reporters for each of three stages of viral replication. These viruses provide a high signal-to-noise ratio while retaining stage specific expression patterns, enabling plate-based assays and microscopic observations of virus propagation and replication. These tools have uses for antiviral discovery, studies of the virus-host interaction, and evolutionary biology.

We describe the usage of a fluorescent reporter vaccinia virus that enables real-time measurement of viral infectivity and gene expression through the stage-specific expression of spectrally distinct reporter fluorophores. We detail a plate-based method for accurately identifying the stage at which virus replication is affected in response to small molecule inhibition.

Poxviruses are a family of double stranded DNA viruses that include active human pathogens such as monkeypox, molluscum contagiousum, and Contagalo virus. ...

Measuring the Kinetics of mRNA Transcription in Single Living Cells

The transcriptional activity of RNA polymerase II (Pol II) is a dynamic process and therefore measuring the kinetics of the transcriptional process in vivo is of importance. Pol II kinetics have been measured using biochemical or molecular methods.1-3 In recent years, with the development of new visualization methods, it has become possible to follow transcription as it occurs in real time in single living cells.4 Herein we describe how to perform analysis of Pol II elongation kinetics on a specific gene in living cells.5, 6 Using a cell line in which a specific gene locus (DNA), its mRNA product, and the final protein product can be fluorescently labeled and visualized in vivo, it is possible to detect the actual transcription of mRNAs on the gene of interest.7, 8 The mRNA is fluorescently tagged using the MS2 system for tagging mRNAs in vivo, where the 3'UTR of the mRNA transcripts contain 24 MS2 stem-loop repeats, which provide highly specific binding sites for the YFP-MS2 coat protein that labels the mRNA as it is transcribed.9 To monitor the kinetics of transcription we use the Fluorescence Recovery After Photobleaching (FRAP) method. By photobleaching the YFP-MS2-tagged nascent transcripts at the site of transcription and then following the recovery of this signal over time, we obtain the synthesis rate of the newly made mRNAs.5 In other words, YFP-MS2 fluorescence recovery reflects the generation of new MS2 stem-loops in the nascent transcripts and their binding by fluorescent free YFP-MS2 molecules entering from the surrounding nucleoplasm. The FRAP recovery curves are then analyzed using mathematical mechanistic models formalized by a series of differential equations, in order to retrieve the kinetic time parameters of transcription.

RNA polymerase II transcriptional kinetics are measured on specific genes in living cells. mRNAs transcribed from the gene of interest are fluorescently tagged and using Fluorescence Recovery After Photobleaching (FRAP) the in vivo kinetics of transcriptional elongation are obtained.

The transcriptional activity of RNA polymerase II (Pol II) is a dynamic process and therefore measuring the kinetics of the transcriptional process in ...

Temporal Analysis of the Nuclear-to-cytoplasmic Translocation of a Herpes Simplex Virus 1 Protein by Immunofluorescent Confocal Microscopy

Infected cell protein 0 (ICP0) of herpes simplex virus 1 (HSV-1) is an immediate early protein containing a RING-type E3 ubiquitin ligase. It is responsible for the proteasomal degradation of host restrictive factors and the subsequent viral gene activation. ICP0 contains a canonical nuclear localization sequence (NLS). It enters the nucleus immediately after de novo synthesis and executes its anti-host defense functions mainly in the nucleus. However, later in infection, ICP0 is found solely in the cytoplasm, suggesting the occurrence of a nuclear-to-cytoplasmic translocation during HSV-1 infection. Presumably ICP0 translocation enables ICP0 to modulate its functions according to its subcellular locations at different infection phases. In order to delineate the biological function and regulatory mechanism of ICP0 nuclear-to-cytoplasmic translocation, we modified an immunofluorescent microscopy method to monitor ICP0 trafficking during HSV-1 infection. This protocol involves immunofluorescent staining, confocal microscope imaging, and nuclear vs. cytoplasmic distribution analysis. The goal of this protocol is to adapt the steady state confocal images taken in a time course into a quantitative documentation of ICP0 movement throughout the lytic infection. We propose that this method can be generalized to quantitatively analyze nuclear vs. cytoplasmic localization of other viral or cellular proteins without involving live imaging technology.

ICP0 undergoes nuclear-to-cytoplasmic translocation during HSV-1 infection. The molecular mechanism of this event is not known. Here we describe the use of confocal microscope as a tool to quantify ICP0 movement in HSV-1 infection, which lays the groundwork for quantitatively analyzing ICP0 translocation in future mechanistic studies.

Infected cell protein 0 (ICP0) of herpes simplex virus 1 (HSV-1) is an immediate early protein containing a RING-type E3 ubiquitin ligase. It is ...

Detection of Viral RNA by Fluorescence in situ Hybridization (FISH)

Viruses that infect cells elicit specific changes to normal cell functions which serve to divert energy and resources for viral replication. Many aspects of host cell function are commandeered by viruses, usually by the expression of viral gene products that recruit host cell proteins and machineries. Moreover, viruses engineer specific membrane organelles or tag on to mobile vesicles and motor proteins to target regions of the cell (during de novo infection, viruses co-opt molecular motor proteins to target the nucleus; later, during virus assembly, they will hijack cellular machineries that will help in the assembly of viruses). Less is understood on how viruses, in particular those with RNA genomes, coordinate the intracellular trafficking of both protein and RNA components and how they achieve assembly of infectious particles at specific loci in the cell. The study of RNA localization began in earlier work. Developing lower eukaryotic embryos and neuronal cells provided important biological information, and also underscored the importance of RNA localization in the programming of gene expression cascades. The study in other organisms and cell systems has yielded similar important information. Viruses are obligate parasites and must utilise their host cells to replicate. Thus, it is critical to understand how RNA viruses direct their RNA genomes from the nucleus, through the nuclear pore, through the cytoplasm and on to one of its final destinations, into progeny virus particles 1.
FISH serves as a useful tool to identify changes in steady-state localization of viral RNA. When combined with immunofluorescence (IF) analysis 22, FISH/IF co-analyses will provide information on the co-localization of proteins with the viral RNA3. This analysis therefore provides a good starting point to test for RNA-protein interactions by other biochemical or biophysical tests 4,5, since co-localization by itself is not enough evidence to be certain of an interaction. In studying viral RNA localization using a method like this, abundant information has been gained on both viral and cellular RNA trafficking events 6. For instance, HIV-1 produces RNA in the nucleus of infected cells but the RNA is only translated in the cytoplasm. When one key viral protein is missing (Rev) 7, FISH of the viral RNA has revealed that the block to viral replication is due to the retention of the HIV-1 genomic RNA in the nucleus 8. 
Here, we present the method for visual analysis of viral genomic RNA in situ. The method makes use of a labelled RNA probe. This probe is designed to be complementary to the viral genomic RNA. During the in vitro synthesis of the antisense RNA probe, the ribonucleotide that is modified with digoxigenin (DIG) is included in an in vitro transcription reaction. Once the probe has hybridized to the target mRNA in cells, subsequent antibody labelling steps (Figure 1) will reveal the localization of the mRNA as well as proteins of interest when performing FISH/IF.

Detection of Viral RNA by Fluorescence in situ Hybridization FISH

A fluorescence in situ hybridization (FISH) method was developed to visually detect viral genomic RNA using fluorescence microscopy. A probe is made with specificity to the viral RNA that can then be identified using a combination of hybridization and immunofluorescence techniques. This technique offers the advantage of identifying the localization of the viral RNA or DNA at steady-state, providing information on the control of intracellular virus trafficking events.

Viruses  that infect cells elicit specific changes to normal cell functions which serve  to divert energy and resources for viral replication. Many ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

Single-Cell Multiplexed Fluorescence Imaging to Visualize Viral Nucleic Acids and Proteins and Monitor HIV, HTLV, HBV, HCV, Zika Virus, and Influenza Infection

Capturing the dynamic replication and assembly processes of viruses has been hindered by the lack of robust in situ hybridization (ISH) technologies that enable sensitive and simultaneous labeling of viral nucleic acid and protein. Conventional DNA fluorescence in situ hybridization (FISH) methods are often not compatible with immunostaining. We have therefore developed an imaging approach, MICDDRP (multiplex immunofluorescent cell-based detection of DNA, RNA and protein), which enables simultaneous single-cell visualization of DNA, RNA, and protein. Compared to conventional DNA FISH, MICDDRP utilizes branched DNA (bDNA) ISH technology, which dramatically improves oligonucleotide probe sensitivity and detection. Small modifications of MICDDRP enable imaging of viral proteins concomitantly with nucleic acids (RNA or DNA) of different strandedness. We have applied these protocols to study the life cycles of multiple viral pathogens, including human immunodeficiency virus (HIV)-1, human T-lymphotropic virus (HTLV)-1, hepatitis B virus (HBV), hepatitis C virus (HCV), Zika virus (ZKV), and influenza A virus (IAV). We demonstrated that we can efficiently label viral nucleic acids and proteins across a diverse range of viruses. These studies can provide us with improved mechanistic understanding of multiple viral systems, and in addition, serve as a template for application of multiplexed fluorescence imaging of DNA, RNA, and protein across a broad spectrum of cellular systems.

Presented here is a protocol for a fluorescence imaging approach, multiplex immunofluorescent cell-based detection of DNA, RNA, and protein (MICDDRP), a method capable of simultaneous fluorescence single-cell visualization of viral protein and nucleic acids of different type and strandedness. This approach can be applied to a diverse range of systems.

Capturing the dynamic replication and assembly processes of viruses has been hindered by the lack of robust in situ hybridization (ISH) technologies that ...